Significance

Neurons typically regulate their intrinsic excitability to prevent excessive excitation and to gate information transfer. This paper describes an activity-dependent decrease in intrinsic excitability following brief bursts of nerve impulses. This homeostatic mechanism, due to the recruitment, or sensitization, of voltage-gated potassium channels, the KCNQ2/3 channels, is reduced in striatal neurons of two transgenic mouse models of Huntington’s disease at an age when these neurons are hyperactive and motor symptoms begin to appear. Pharmacological activation of these channels restores homeostasis in transgenic neurons, in vitro, and reduces motor impairment in behaving mice, consistent with the hypothesis that hyperactivity enables establishment of dysfunctional neural circuits and that KCNQ channels could serve as therapeutic targets for the treatment of HD.

Abstract

We describe a fast activity-dependent homeostatic regulation of intrinsic excitability of identified neurons in mouse dorsal striatum, the striatal output neurons. It can be induced by brief bursts of activity, is expressed on a time scale of seconds, limits repetitive firing, and can convert regular firing patterns to irregular ones. We show it is due to progressive recruitment of the KCNQ2/3 channels that generate the M current. This homeostatic mechanism is significantly reduced in striatal output neurons of the R6/2 transgenic mouse model of Huntington’s disease, at an age when the neurons are hyperactive in vivo and the mice begin to exhibit locomotor impairment. Furthermore, it can be rescued by bath perfusion with retigabine, a KCNQ channel activator, and chronic treatment improves locomotor performance. Thus, M-current dysfunction may contribute to the hyperactivity and network dysregulation characteristic of this neurodegenerative disease, and KCNQ2/3 channel regulation may be a target for therapeutic intervention.

Huntington’s disease (HD) is a fatal inherited autosomal neurodegenerative disorder, with its primary symptoms being progressive development of motor and cognitive dysfunction (1). The mutated gene, huntingtin (HTT), and its mutation, an expansion of the number of CAG repeats, were identified 20 y ago. However, the mechanism(s) underlying the pathological changes that culminate in the degeneration of striatal output neurons (SONs) remain unknown. Early animal models (2) generated a number of testable hypotheses, most notable being that the neurons degenerate because of a hyperactivity that leads to a build-up of excitotoxic molecules. However, more recent studies implicate alternative pathologies, such as altered transcriptional activity, calcium regulation and mitochondrial function, or disruptions in normal neuronal patterns of activity (3) and show that neuronal dysfunction and behavioral and motor symptoms of HD precede neurodegeneration (2). These studies have been facilitated by access to transgenic mice models, including R6/1 and R6/2 mice, which express a truncated region of the mutant human HTT gene with expanded CAG repeats (4). In vitro recordings in both lines revealed that SONs are depolarized and have higher input resistances than do wild-type (WT) controls, at a stage where deficits in locomotor activity begin to be manifest (5⇓–7). Furthermore, in vivo recordings indicate that at 5–9 wk of age, when the mice exhibit overt motor deficits, R6/2 SONs have higher firing rates and more regular discharge patterns compared with WT (8, 9). In contrast, neurodegeneration and death occur later (2). Hence, we asked whether cellular mechanisms that influence excitability might be altered in the early stages of HD and might serve as targets for alleviating associated behavioral symptoms.

Hyperactivity and related changes in neuronal firing patterns could reflect alterations in synaptic transmission and its activity-dependent modifications or in intrinsic membrane properties governing neuronal excitability (10⇓–12). The latter can also be modulated by activity (13) and have homeostatic roles (14). We describe here a fast activity-dependent homeostatic control of excitability (fADH) in SONs. In WT mice, fADH can be induced by brief trains of impulses and is expressed on a time scale of seconds. It modifies firing rate and timing of evoked spikes, converting regular firing patterns to irregular ones, with the latter mode resembling the accommodation attributed to voltage- and time-dependent activation of the M current mediated by KCNQ [or voltage-gated potassium channel (Kv) subfamily 7 or Kv7] channels (14). Indeed, increasing activation of KCNQ channels on successive trials underlies fADH. Strikingly, we found that fADH is reduced in R6/2 SONs, that two KCNQ activators (15, 16) rescued fADH in R6/2 SONs, thereby restoring WT firing patterns, and that the locomotor signs of HD in the R6/2 mouse were ameliorated by chronic treatment with one of the activators.

Results

Basic Membrane Properties of WT and R6/2 SONs.

Comparison of SON basic membrane properties for 4- to 6-wk-old mice revealed an increased input resistance in the transgenics, from 43.7 ± 3.8 MΩ to 68.2 ± 7.3 MΩ, with a corresponding decrease in rheobase, the threshold current for a long pulse, from 201.3 ± 15.4 pA to 138.3 ± 13.0 pA, and without a detectable change in the threshold voltage (Table S1). Comparable results were found when the data analysis was restricted to recordings with a series resistance < 25 MΩ, although the decrease in rheobase did not reach significance with the smaller sample sizes (Table S1). These observations are similar to those reported for R6/2 SONs at an overlapping range of ages, 5–7 wk (5, 17). However, there was no difference in resting membrane potential (Table S1), in contrast to the depolarization reported for the slightly older R6/2 neurons. Because WT SONs tend to discharge in brief bursts in vivo (18, 19), we asked whether the relationship between stimulus current (I) and impulse frequency (f) was altered, using 300-ms suprathreshold depolarizing pulses to evoke action potential trains, with a stimulus frequency (<0.5 Hz) that does not induce fADH. The f-I relations were essentially the same, after normalizing stimulus strength with respect to rheobase, and the cells had the same maximum discharge rates (Fig. 1A). Thus, a change in input resistance per se would not necessarily account for differences in firing patterns observed in vivo.

Impairment of fADH in R6/2 SONs. (A) WT and R6/2 SONs have similar f-I relations and asymptotic firing rates. Plot of mean impulse frequency during a 300-ms depolarizing current pulse versus current amplitude, normalized with respect to rheobase. Also shown are exponential fits of data from four WT (black) and seven R6/2 (red) SONs. (B) fADH is prominent in WT (Left) but not in R6/2 (Right) SONs. Top three traces in each column are first, 10th and 20th spike trains evoked by the current pulse repeated at 1 Hz. Stimulus strength, 2× rheobase. Bottom traces, recovery after 1-min rest and sample current pulse. (C and D) Characterization of fADH. (C) Cumulative distributions of I20 for WT (black) and R6/2 (red) SONs (n = 62), MW U test, ***P < 0.001, KS test, ***P < 0.001. (D) Progressive build-up of adaptation, and recovery kinetics, plotted as Adaptation Index (In, ordinate), versus time (abscissa). Recovery data obtained by following conditioning series with rest periods of 2, 5, 10, 30, and 60 s (one test pulse per conditioning series). Time constants of development, τdev, and recovery, τrec, calculated from monoexponential fits of pooled data (7 SONs).

Characterization of fADH.

We noted that the responsiveness of WT SONs to a suprathreshold 300-ms depolarizing pulse repeated at 1-s intervals gradually became weaker; the stimulus initially evoked a train of ∼10–13 impulses (Fig. 1B), but the firing frequency steadily decreased with successive stimuli and was reduced by ∼50% after 10–20 trials. Fig. 1B contrasts this fADH in the WT with the minimal accommodation, or adaptation, exhibited by R6/2 SONs. For quantification, we defined the Adaptation Index, In = 1 − (no. of spikes, nth trial/no. of spikes, first trial); theoretically, In ranges from 0 to 1.0 (maximal adaptation to no evoked spikes). In Fig. 1B, for the WT SON, I10 and I20 = 0.25 and 0.50, respectively, whereas for the R6/2 example, both indices = 0.14. Overall, the transgenics exhibited a significant shift to the left in the cumulative distribution for I20 (Fig. 1C), which averaged 0.26 ± 0.02, compared with 0.49 ± 0.03 (n = 62 each) for the WT neurons. The differences between the medians and between the distributions are all highly significant (P < 0.001).

There are two populations of SONs, those expressing D1- or D2-dopamine receptors, also known as the direct and indirect pathway neurons, with reported differences in electrophysiological properties, namely that D1 SONs apparently have a higher rheobase and a lower input resistance and fire at lower frequencies to the same magnitude depolarizing current pulse (20). These correlations were characteristic of our results, as shown by separately sorting the WT and R6/2 datasets into subpopulations defined by those neurons with high and low rheobase values (upper and lower 25% or 50%), and there were no differences in I20 (Fig. S1A). In contrast, when the data were sorted according to high or low values of I20, rheobase, input resistance, and evoked spike rate were the same, in WT or in R6/2 (Fig. S1B). These findings, coupled with the unimodal character of the cumulative distributions for I20, suggest fADH is expressed in both neuronal subtypes in WT and is diminished in both in the HD model.

fADH was manifest within seconds and was cumulative. With the standard protocol, it developed with a time constant of ∼10.5 s and decayed with a time constant of 16.3 s, such that recovery was complete with 1- to 2-min rest (Fig. 1 B and D). It was not associated with changes in input resistance, spike height or half-width, as measured for the first spike in an evoked train, or resting membrane potential. Also, it could be induced with other conditioning frequencies, in the range of 0.5–2.0 Hz, with a tendency to increase as a function of both stimulus strength and frequency (Fig. 2A). For example, on average, I20 increased by 78% and 83% when conditioning frequency was doubled from 1 to 2 Hz while maintaining the strength constant at 2.0 or 2.8 times rheobase, respectively, and by 28% and 31% when the strength was increased while holding frequency constant at 1 or 2 Hz, respectively. These results suggest the magnitude of fADH depends on the time integral of the conditioning depolarization; however, subthreshold depolarizations did not induce fADH. Finally, the magnitude of the Adaptation Index grew progressively as a function of the cumulative number of spike discharges in the preceding 20 s (Fig. 2B), as might be expected of a spike-counting mechanism characterized by interplay between activation and recovery processes with similar kinetics.

Characterization of fADH. (A) Adaptation Indices after 10, 20, and 30 trials, plotted as functions of stimulus frequency (1 and 2 Hz) and strength (2, 2.4, 2.8, and 3.2 times rheobase, Rb). (B) Adaptation Index versus cumulative number of spikes in the preceding 20 trials (n = 4 experiments) with exponential fit (red). (C1, C2, and D) Influence of fADH on spike firing patterns. (C1 and C2) Spike raster plots from the experiments of Fig. 1B, contrasting expression of fADH in WT (1) with that in R6/2 (2). Timing of successive evoked spikes in each train is marked by color-coded symbols, with trial number increasing from top to bottom. (D) Deficit in fADH in R6/2 minimizes variability in spike timing. Mean interspike interval (ISI) for the first and fourth intervals in the trains and mean first spike latency plotted versus trace number for WT (black) and R6/2 (red) (7 WT and 14 R6/2 SONs). (E1 and E2) fADH evoked by brief stimulus trains. Same format as in Fig. 1B but stimulation was with a train of 10 suprathreshold pulses (2x Rb, 10-ms duration, 30-ms cycle time). Evoked spike train in R6/2 (2) is at a higher frequency and is more regular than that in WT (1).

We asked whether fADH was a consequence of dialysis of the neuronal cytoplasm, by recording first in the cell attached mode and stimulating extracellularly through the same electrode, allowing direct comparison with subsequent recordings from the same neuron in whole cell mode. fADH was observed in both modes, with the Adaptation Index being greater, on average, in the latter (Fig. S2A). Also, the magnitude of fADH was the same after warming the slices from 26.0 ± 0.6 to 30.7 ± 0.4 °C, suggesting it is operational at physiological temperatures (Fig. S2B).

fADH Alters Structure of Impulse Trains.

fADH introduces both a progressive slowing of impulse activity within an evoked burst of spikes and an increased variability in the interspike interval, whereas impulse trains in R6/2 SONs are more regular and predictable (Fig. 2 C1, C2, and D). It could be evoked with other stimulating paradigms, such as ones that mimicked repetitive synaptic input, and in that case, the evoked discharge pattern became irregular in WT but not in the transgenic neurons (Fig. 2 E1 and E2). Thus, the stimulus protocols used here evoked discharge patterns similar to those of WT SONs recorded in vivo at a similar age, namely an overall low frequency firing rate characterized by brief bursts separated by periods of quiescence (18, 19). In contrast, R6/2 SONs discharge at higher frequencies and are more regular (9), consistent with the hypothesis that these differences reflect the differential expression of fADH. Therefore, we investigated the mechanism underlying fADH.

fADH Is Due to Increased M-Current Activation.

Potential mechanisms underlying fADH include Ca2+- and voltage-dependent modulation of ion channels that shape neuronal activity patterns, such as a persistent subthreshold Na+ current (21) or various K+ channels (11, 22). Because neuronal activity is typically associated with an increase in intracellular [Ca2+], we first asked whether chelating this cation with 25 mM BAPTA in the recording pipette affected fADH. However, BAPTA had no effect on fADH, as seen by comparing the cumulative distributions of I20 in its presence or absence (Fig. S3). We then focused on K+ channels, specifically the KCNQ channels that generate the voltage-dependent M-current that contributes to accommodation in a number of cell types (23, 24) and is expressed by SONs (25).

The KCNQ channel blocker XE-991 (6 μM) was bath-applied while recording from SONs, and it significantly reduced adaptation (Fig. 3A). In the illustrated case, I20 decreased from 0.6 to 0.18. The mean adaptation indices I10 and I20 from 7 SONs (31–37 d postnatal) in which the drug effect was followed by successful wash out (recovery to within 10% of control) were both reduced by 50% or more (Fig. 3C).

fADH is due to activity-dependent enhancement of M current. (A) fADH in control (Left) and in the presence of 6 μM XE-991 (Right). Traces are, from top to bottom, first and 20th trials, and recovery. Stimulus: 1.8x Rb. (B) Voltage clamp recordings of current responses to a 70-mV step from −90 mV, in control (Upper) and after adding XE-991 (Middle). Responses obtained before (black) and after (red) inducing fADH. (Lower) XE-991 sensitive currents in the two conditions. (C) Bar plots of effects of 6 μM XE-991 (Upper), 10 μM McN-A-343 (Middle), and 10 μM Wortmannin (Lower), on I10 and I20. n = 7, 7, and 5, for Control, XE-991 and washout, respectively; n = 8, 8, and 5, for Control, McN-A-343, and washout, respectively; n = 6, 6, and 4, for Control, Wortmannin, and washout, respectively; error bars are SEM; Student’s t test and Welch’s t test, not significant (n.s.) P > 0.05, *P < 0.05, **P < 0.01. (D and E) Same format as in A, but for 10 μM McN-A-343 and 10 μM Wortmannin, respectively.

Finally, Fig. 3B demonstrates directly, with voltage-clamp recordings, that the induction protocol and fADH are associated with an enhanced outward current response to a 70-mV, 300-ms depolarizing step from −90 mV (Upper) and that in the presence of XE-991, there is no additional current induced by the conditioning paradigm (Middle). The magnitude of the XE-991 sensitive current is increased by approximately 60% after induction of fADH (Lower). In contrast, XE-991 had no effect on the responses of the R6/2 neurons. An alternative method for quantifying M current is as the current that deactivates when membrane potential is stepped back from −20 to −50 mV (25). The amplitude of the deactivating current was enhanced after fADH induction, and this increase decayed back toward baseline over a period of 60–90 s, comparable to the time course of the decay of fADH (Fig. S4). Also, comparison of normalized I-V plots before conditioning did not reveal any differences in voltage dependence of M-current activation between WT and R6/2 SONs (Fig. S4).

The M-current magnitude is sensitive to the level of phosphatidylinositol 4,5-bisphosphate (PIP2) in the plasma membrane, through mechanisms that may alter channel activation, for instance, by regulating channel open probability, voltage dependence, or number (26). PIP2 hydrolysis is increased by activation of M1 muscarinic receptors, through a G protein-coupled receptor mediated cascade, and SON excitability is enhanced by cholinergic inhibition of KCNQ channels (26). As predicted, the M1 agonist, McN-A-343 (10 μM) reversibly reduced fADH (Fig. 3 C and D). In the illustrated example, I20 was reduced by 50%, and overall I10 and I20 were reduced by 47 and 40%, respectively, from 0.38 ± 0.04 to 0.20 ± 0.03 (P < 0.01) and from 0.42 ± 0.04 to 0.25 ± 0.03 (P < 0.01, n = 8; in 5 experiments, the block was reversed by washout). Also, the activity of PI-4 kinase, which catalyzes the synthesis of PIP in the plasma membrane, can be enhanced by depolarization, favoring increased downstream synthesis of PIP2 (27⇓–29). This first step in the synthetic cascade can be blocked by high concentrations (10 μM) of Wortmannin, and indeed, this compound significantly reduces fADH (Fig. 3 C and E). Taken together, these results are consistent with the hypothesis that fADH is due to activity-dependent sensitization of the M current.

We performed Western blot analysis of KCNQ2 expression in dorsal striatum and prefrontal cortex of 5-wk-old WT and R6/2 mice. Expression of two inward rectifying channel proteins, Kir2.1 and Kir2.3 (30), was also quantified. Although KCNQ2 abundance in cortex appeared to be reduced in the transgenic, the effect was not significant, and we found no significant difference in its level in striatum (Fig. S5). Also, Kir2.1 expression was reduced only in striatum, whereas Kir2.3 was reduced in both structures. The maintained expression of KCNQ2 is consistent with the demonstration that these channels can be activated in R6/2 SONs, as described below.

M-Current Activators Rescue fADH in R6/2 Neurons.

We next tested the possibility that the reduction in fADH in R6/2 SONs reflects altered channel regulation by asking whether M-current activators could reverse this functional deficit. Fig. 4 A and C and Fig. 4 B and D illustrate results obtained from R6/2 SONs during bath superfusion of 25 μM diclofenac (15) and 10 μM retigabine (16), respectively. In both, the activators, which shift the conductance-voltage relation to the left on the voltage axis, reversibly enabled fADH, increasing I20 by a factor of 2–3. On average, diclofenac increased I20 from 0.22 ± 0.02 to 0.50 ± 0.06 (n = 8, P < 0.01) whereas the control and washout data were statistically the same (P > 0.45). The corresponding values for retigabine are 0.24 ± 0.03 vs. 0.54 ± 0.05 (P < 0.001). This effect is in contrast to the observation that the magnitude of fADH in WT SONs was unaffected by retigabine (Fig. S6). Finally, increasing intracellular PIP2 concentration by including diC8-PIP2 (31) in the recording pipette solution also rescued fADH in R6/2 SONs (Fig. S6). These findings indicate that the transgenic SONs have latent activatable KCNQ channels.

Multiple transgenic mouse models are used to study HD, and the temporal progression of the signs and symptoms differ, depending on the construct and the number of CAG repeats. For example, motor signs seen at 4 wk of age in the truncated R6/2 model (32), appear later, at approximately 2 mo, in the BACHD mouse, which has the full-length mutated human gene (33, 34). We thus asked whether fADH could be induced in the WT mice used to generate the BACHD and whether there is a deficit in this adaptive mechanism at 2 mo in the BACHD mice. SONs from both sets of mice exhibited fADH, and it was quantitatively smaller in the BACHD SONs (Fig. S7). Specifically, the mean Adaptation Index was decreased by 34% and 31% after 10 and 20 trials, respectively. This result indicates there is a similar deficit in fADH at quite different ages, but at a comparable stage of the pathology. We also confirmed that fADH could be rescued in the affected BACHD neurons by applying retigabine; in 8 SONs, it increased I10 from 0.13 ± 0.019 to 0.24 ± 0.044 (P < 0.05) and I20 from 0.19 ± 0.02 to 0.31 ± 0.052 (P < 0.05).

As noted, R6/2 mice begin to exhibit locomotor deficits as early as 4–5 wk of age (32). Because our electrophysiological results indicate there is a deficit in fADH at the same age, we asked whether chronic treatment with retigabine would reduce or slow the development of these deficits, as measured by performance in the open field test, and positive results were obtained across a 3-wk testing period (Fig. 5). Mice were treated with daily i.p. injections of retigabine (10 mg/kg), starting at 4 wk of age and were tested once per week from week 5 through week 7. In this age range, the R6/2 controls did not exhibit seizures, and thus any effects of retigabine would not be due to its antiepileptic properties. General locomotor activity was assessed as track length, and exploration as the number of rearings, in a 9-min period. For both measures, the WT controls that were untreated, vehicle-treated, and drug-treated were statistically equivalent, and the first two groups are combined in the analysis and graphs. The same was the case for the R6/2 untreated and vehicle-treated controls, and they are referred to as control R6/2. The figure illustrates that the control R6/2 mice are significantly impaired in both measures of general activity and exploration, compared with WT, across the 3-wk testing period. More importantly, administration of retigabine significantly improved locomotor activity in treated R6/2 mice compared with control R6/2 mice across the 3-wk test period. The data also demonstrate a trend, albeit not statistically significant, toward improvement in the number of rearings in treated R6/2 mice compared with control R6/2 mice. Thus, the M-current activator rescued both electrophysiological and behavioral phenotypes of the transgenic HD model. Finally, because the age of the mice used for the behavioral studies encompassed 5–7 wk, we confirmed that fADH was still differentially expressed in WT and R6/2 at 7–8 wk (Fig. S8).

Discussion

The major findings of this study are that SONs exhibit activity-dependent homeostatic regulation of excitability that is associated with enhanced activation of the M current (11) and is depressed in SONs in two transgenic mouse models of HD, at ages when motor symptoms are just emerging (32⇓–34). Moreover, this deficit is at least partially reversed by acute exposure to M-current activators. Finally, daily treatment of the R6/2 mice with an activator improves motor performance in the affected mice.

We propose that fADH is due to a progressive activity-dependent enhanced activation, or sensitization, of KCNQ channels, such that after one brief train of impulses, the next excitation can generate a larger K+ conductance and slow neuronal spiking. Presumably, deactivation of the conductance between test stimuli and the relatively slow time constant of activation of the M current by depolarization, 80–170 ms, in SONs (25) and in CHO cells (35), explains why this modulation has a minimal effect on input resistance or the timing of the first impulse within a burst. We suggest that the observed increase in the outward XE-991–sensitive currents after fADH induction most likely reflects a shift in open probability or an increase in the number of available channels (27), consistent with the effect of PIP2. Although a shift in the voltage dependence of the channels, as occurs with retigabine (16), cannot be ruled out, it is less likely, given that the normalized I-V plots of WT and R6/2 SONs are the same for depolarizations up to −20 mV. Regardless, our results are consistent with the previous demonstration that M-current suppression increases SON excitability (25). The increased input resistance, which could reflect down-regulation of a second channel, would also contribute to increased excitability.

The temporal domain of fADH is suggestive of real-time adjustments in intrinsic excitability, and distinguishes it from slower developing long-term forms of intrinsic plasticity, which also reflect K+ channel regulation (13, 14). Thus, fADH is a form of metaplasticity, and it is due to time-dependent M-current recruitment in response to prolonged or repeated excitation, presumably by increasing the level of PIP2 in the membrane, and it can be expected to have a braking effect on neuronal activity.

The effects of Wortmannin and PIP2 are consistent with the hypothesis that fADH is the consequence of a depolarization-induced increased synthesis of PIP2, through a membrane-bound enzymatic cascade. Huntingtin inserts in the plasma membrane through interactions with phosphoinositides, and N-terminal huntingtin fragments target membrane regions enriched in PIP2 (36). Furthermore, these interactions are altered by polyglutamine expansions of the N-terminal fragments, including a shift in targeting toward intracellular regions (37, 38). This aberrant molecular organization might well underlie the deficit in fADH in the HD mice.

A slower and more persistent form of activity-dependent homeostatic intrinsic plasticity, exhibited by hippocampal pyramidal neurons, is also mediated by KCNQ channels (14). However, in that case, the induction mechanism is Ca2+-dependent, leading to the suggestion that complementary homeostatic mechanisms operating through separate but convergent signaling pathways may function to place an upper limit on neuronal firing, and to guarantee a degree of variability in impulse pattern.

There is a striking agreement among the ages at which R6/2 mice first become symptomatic, demonstrate increased activity and altered firing patterns in vivo, and exhibit a deficit in fADH. This correlation is strengthened by the parallel observations with the BACHD mice. In addition, evidence that KCNQ channels can be recruited in the transgenic neurons, coupled with the effects, at the same age, of the M-current activator retigabine on motor performance, identifies a potential for reducing both neuronal hyperactivity and motor impairment. Its effect on locomotor activity presumably is not due to a nonspecific reduction in neuronal activity, because it had no effect on WT mice.

Homeostatic mechanisms can have significant effects on network development and function (39, 40). As one example, suppression of KCNQ2 expression increases the excitability and reduces spike-frequency adaptation of hippocampal CA1 pyramidal cells and alters hippocampal morphology, with associated behavioral changes, and the manifestation of these effects depends on the time window of suppression (41). Given these considerations, we suggest that fADH may have a critical role in the formation and stability of the circuitry in dorsal striatum, for example, by preventing the aberrant hyperactivity that has been implicated in the etiology of a number of neurodegenerative disorders (42).

Experimental Procedures

Electrophysiology.

Parasagittal brain slices (300 μm thick) were prepared, as described (43), from transgenic R6/2 mice and their WT (CBA × C57BL/6J) littermates (postnatal days 25–57) as described (43), and from BACHD mice and their WT littermates (FVB/N). Animals were bred and genotyped at Jackson Labs and provided by the CHDI Foundation. Animal handling and use followed a protocol approved by the Institutional Animal Care and Use Committee of Albert Einstein College of Medicine. Briefly, animals were decapitated under isofluorane anesthesia. The brain was removed quickly and transferred into ice-cold saline that contained the following (in mM): 125 NaCl, 4 KCl, 10 glucose, 1.25 NaH2PO4, 25 NaHCO3, 0.5 CaCl2, and 2.5 MgCl2, equilibrated with a 5% CO2-95% O2 mixture (pH 7.3). The two sagittal brain hemispheres were separated and adhered to the stage of a Vibratome Series 1000 Classic vibroslicer, and slices were cut at an angle of 10 ± 2° (44). Slices were allowed to recover for at least 1 h at room temperature in artificial cerebrospinal fluid (ACSF) consisting of (in mM): 125 NaCl, 4 KCl, 10 glucose, 1.25 NaH2PO4, 25 NaHCO3, 2 CaCl2 and 1 MgCl2 (pH buffered to 7.3 with 5% CO2-95% O2).

Whole-cell voltage- and current-clamp recordings were performed, using IR-differential interference contrast (DIC) video microscopy to visualize and photograph individual neurons, and cell types were distinguished on the basis of characteristic morphology, action potential waveforms and responses to 300–400 ms depolarizing and hyperpolarizing current pulses. To quantify input resistance, we used the slope of first part in the hyperpolarizing quadrant of the I-V function for small hyperpolarizing and depolarizing current pulses, thereby avoiding the region of pronounced inward rectification.

Recordings were at room temperature, unless stated otherwise, with patch-type pipette electrodes (4–8 MΩ) filled with (in mM) 100 potassium gluconate, 50 KCl, 5 NaCl, 0.5 CaCl2, 5 EGTA, 25 Hepes, 2 MgATP, 0.3 GTP (pH 7.2). Series resistance (<40 MΩ) was monitored throughout each experiment, and data from cells with more than 20% changes in series resistance were discarded. Although 40 MΩ is commonly used as a cutoff in recordings from SONs (14, 25), when the data analysis was restricted to values <25 MΩ, results were essentially the same (Table S1). Electrophysiological data were digitized at 20 kHz with a National Instruments BNC 2090 A-D convertor connected to a Macintosh personal computer and analyzed with customized software, as well as in IGOR and Origin. All data are from neurons with resting membrane potentials of magnitude greater than −70 mV.

Statistical Analysis.

Values are expressed as mean ± SEM (n = number of cells). Two-tailed Student’s and Welch’s t tests were used for statistical comparisons at the P < 0.05 significance level. Cumulative probability plots were compared by using the Kolmogorov–Smirnov (KS) test and the Mann–Whitney (MW) U test. Behavioral data were analyzed by using a repeated-measures ANOVA.

Drugs.

XE-991 and Wortmannin (Ascent Scientific), diclofenac and McN-A-343 (Sigma-Aldrich), and retigabine (provided by CHDI Foundation) were added to ACSF after obtaining control data, and at least 15 min was allowed to lapse before recording in the presence of the drug. To test for involvement of activity-dependent rises in intracellular Ca2+ concentration, 25 mM BAPTA was added directly to the pipette solution. To test the PIP2 hypothesis, cells were loaded with a water soluble form of PIP2, diC8-PIP2 (Cayman Chemical) via the patch pipette. diC8-PIP2 was dissolved in deionized water at 1 μg/mL concentration and stored in aliquots at −20 °C (31). An aliquot was thawed, diluted in internal solution at 30 μM final concentration, and used immediately.

Open Field.

There were three groups each of WT and R6/2 mice: untreated controls, those treated with 10 mg/kg (i.p.) retigabine daily and those injected with vehicle. Treatment started in week 4 and testing in week 5. Mice were placed in an opaque Plexiglas arena (16 square inches) and allowed to freely explore the arena for 9 min, during which time voluntary locomotion (total distance traveled in centimeters), exploration (number of rears, defined as lifting of the upper body and forepaws off the ground, whisking and sniffing) were scored while being recorded digitally. Longer test durations were not used, because they are often associated with prolonged periods of inactivity, i.e., habituation. Track length was scored automatically with Viewer tracking software (Biobserve) and rears were scored manually. All measurements were done blind to genotype and treatment condition.

Acknowledgments

We thank R. Grantyn for comments on the manuscript and for suggestions about experimental design. This work was supported by CHDI Foundation (A-3674), NIH (NS050808, NS079750, and RR0277888), and Fondecyt (11140430).

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